Hypertrophy in Skeletal Myotubes Induced by Junctophilin-2 Mutant, Y141H, Involves an Increase in Store-operated Ca²⁺ Entry via Orai1

Background: Junctophilin-2 (JP2) contributes to the formation of junctional membrane complexes (JMC) in striated muscle.

Results: Different from the S165F mutant of JP2, Y141H induces hypertrophy in skeletal myotubes involving abnormal JMC and altered Ca²⁺ signaling due to the increased store-operated Ca²⁺ entry (SOCE) via Orai1.

Conclusion: JP2 is linked to muscle hypertrophy via various Ca²⁺ signaling pathways.

Significance: SOCE is a novel factor in understanding muscle hypertrophy.

Abstract

Junctophilins (JPs) play an important role in the formation of junctional membrane complexes (JMC) in striated muscle by physically linking the transverse-tubule and sarcoplasmic reticulum (SR) membranes. Researchers have found five JP2 mutants in humans with hypertrophic cardiomyopathy. Among these, Y141H and S165F are associated with severely altered Ca²⁺ signaling in cardiomyocytes. We previously reported that S165F also induced both hypertrophy and altered intracellular Ca²⁺ signaling in mouse skeletal myotubes. In the present study, we attempted to identify the dominant-negative role(s) of Y141H in primary mouse skeletal myotubes. Consistent with S165F, Y141H led to hypertrophy and altered Ca²⁺ signaling (a decrease in the gain of excitation-contraction coupling and an increase in the resting level of myoplasmic Ca²⁺). However, unlike S165F, neither ryanodine receptor 1-mediated Ca²⁺ release from the SR nor the phosphorylation of the mutated JP2 by protein kinase C was related to the altered Ca²⁺ signaling by Y141H. Instead, abnormal JMC and increased SOCE via Orai1 were found, suggesting that the hypertrophy caused by Y141H progressed differently from S165F. Therefore JP2 can be linked to skeletal muscle hypertrophy via various Ca²⁺ signaling pathways, and SOCE could be one of the causes of altered Ca²⁺ signaling observed in muscle hypertrophy.

Introduction

In striated muscle, the dihydropyridine receptor (DHPR)² (which is a membrane voltage-sensing protein as well as a Ca²⁺-entry channel on transverse (t)-tubule membranes) is activated by membrane depolarization, which allows the ryanodine receptor (RyR, which is an internal Ca²⁺-releasing channel on the sarcoplasmic reticulum (SR) membrane) to release Ca²⁺ ions from the SR into the myoplasm, and finally induces muscle contraction. This process is called excitation-contraction (EC) coupling (1–3). In addition to the central theme of EC coupling, canonical-type transient receptor potential cation channel 3 (TRPC3) (which is a Ca²⁺ entry channel on sarcolemmal membrane) is required for the full gain of skeletal EC coupling (duration and maintenance) (4). Collaterally, channels mediating store-operated Ca²⁺ entry (SOCE) (5, 6) are needed for the sustained presence of a certain concentration of myoplasmic Ca²⁺ ions in skeletal muscle cells: for example, Orai1, TRPC1, and TRPC4 (7–11). Junctional membrane complexes (JMC), which are known as diad or triad junctions in striated muscle and where t-tubules and the SR membranes are juxtaposed, provide the structural context for proper arrangement of the Ca²⁺ channels mentioned above and for functionally efficient couplings among the Ca²⁺ channels (12, 13).

Junctophilins (JPs) contribute to the formation of JMC in excitable cells, including muscle cells, by interacting with both plasma and endoplasmic reticulum membranes (in muscle cells, t-tubule membranes via their N-terminal MORN motifs (highly conserved membrane occupation and recognition nexus) and SR membranes via the C-terminal transmembrane domain) (12–15). Thus far, four JP subtypes have been identified: JP1 in skeletal muscle; JP2 abundantly in all muscle types (skeletal, cardiac, and smooth muscles); and JP3 and JP4 in the brain (14, 16, 17). JP1 or JP2 knock-out in mice induces disorganized JMC and disrupted Ca²⁺ homeostasis in skeletal muscle or cardiomyocytes, and finally neonatal or embryonic death, respectively (12, 13, 18). JP1 and JP2 knockdown in mouse skeletal muscle fibers leads to decreases in intracellular Ca²⁺ release and SOCE and the disorganization of JMC (19). TRPC3 knockdown in mouse skeletal myotubes involves an increase in JP1 expression and a decrease in Ca²⁺ transients from the SR (4), and the N-terminal region of JP2 (amino acids 143–234) interacts with TRPC3 in skeletal myotubes (20, 21). In the case of the neuronal JP isoforms (JP3 and JP4), the only information that has been reported involves a mouse model for neurodegenerative Huntington disease-like 2 disorder that showed a decrease in JP3 expression due to changes in the genetic level (22).

Hypertrophic cardiomyopathy (HCM) in humans, a significant cause of sudden cardiac death, is a genetic disorder related to mutations of muscle genes such as those encoding actin, myosin, tropomyosin, troponin, Z-disc proteins, and others (23). Down-regulation of JP2 has been observed in hypertrophic and dilated cardiomyopathic mouse models (24), in a pressure overload-induced cardiac hypertrophic rat model (25), and over the progression of hypertrophy into heart failure in a rat model (26). Indeed, five JP2 single mutants (S101R, Y141H, S165F, R436C, and G505S) have been found in HCM patients (27, 28), which strongly supports the idea that JP2 is one of the HCM-related proteins. Among the five mutants of JP2, Y141H and S165F significantly disrupt Ca²⁺ signaling when expressed in HL-1 atrial cardiomyocytes (27). In mouse skeletal myotubes, S165F also induced hypertrophy and a reduced gain in EC coupling, which is related to the phosphorylation of S165F by protein kinase C (PKC) (29). However, the effect(s) of the four other human HCM-related JP mutants on skeletal muscle have not been addressed, although JP2 is an abundant protein in both skeletal and cardiac muscles. Therefore, in the present study, Y141H, a JP2 mutant associated with human HCM and one of two mutants causing severely disrupted Ca²⁺ signaling in cardiomyocytes, was expressed in mouse skeletal myotubes to examine the effect(s) of Y141H on skeletal muscle function and Ca²⁺ signaling.

EXPERIMENTAL PROCEDURES

Cell Culture and Expression of Wild-type JP2, Y141H, and S165F

Mouse primary skeletal myoblasts were derived from mouse skeletal muscle as previously described (30). All surgical interventions and presurgical and postsurgical animal care was provided in accordance with the Laboratory Animals Welfare Act, the Guide for the Care and Use of Laboratory Animals and the Guidelines and Policies for Rodent Survival Surgery provided by Institutional Animal Care and Use Committee in the College of Medicine, The Catholic University of Korea, Korea. Primary myoblasts were cultured on 10-cm dishes coated with collagen in growth medium (F10 Nutrient Mixture containing 20% fetal bovine serum (FBS), 100 units/ml of penicillin, 100 μg/ml of streptomycin, 2 mm l-glutamine, and 20 nm basic fibroblast growth factor) at 37 °C in a 5% CO₂ incubator. For differentiation into myotubes, myoblasts were replated on 10-cm dishes (for the myotube lysate preparation or the cell migration assay), 96-well plates (for the single myotube Ca²⁺ imaging experiment or the immunocytochemistry), or 6-well plates (for the TEM observation) coated with Matrigel (BD Biosciences). When myoblasts reached ∼70% confluence, the growth medium was replaced with differentiation medium (5% heat-inactivated horse serum and low-glucose Dulbecco's modified Eagle's medium (DMEM) without growth factors instead of 20% FBS and F-10 Nutrient Mixture in the growth medium), and the myoblasts were placed in an 18% CO₂ incubator to induce differentiation. After 3 days of culture in differentiation medium, the myoblasts were transfected with cDNA encoding wild-type JP2, Y141H, or S165F in an expression vector (pCMS-RFP) for 4 h. Construction of the cDNAs was previously described (27). A mixture of 30 μl of FuGENE 6 (Roche Diagnostics) and 20 μg of cDNA per 10-cm dish or the same ratio of the components in the wells of 96- or 6-well plates was used for transfection. Fully differentiated myotubes on differentiation day 5 (at 36 h post-transfection) were subjected to further experiments. Approximately 40% of the myotubes had been transfected by the cDNA (estimated by a red fluorescent protein (RFP) signal from a separate promoter of the expression vector). For the cell migration assay, premature myotubes at 16 h post-transfection were used.

HL-1 atrial cardiomyocytes were plated on 10-cm dishes coated with fibronectin (Sigma) and gelatin (Fisher Scientific) and maintained in Claycomb media (JRH Biosciences) supplemented with 10% FBS, 1% penicillin/streptomycin, 0.1 mm norepinephrine, and 2 mm l-glutamine. HL-1 cardiomyocytes were transfected with cDNA encoding wild-type JP2 or Y141H in the expression vector as described above. At 36 h post-transfection, HL-1 cardiomyocytes were subjected to the cell lysate preparation or the image capture. Approximately 50% of the HL-1 cardiomyocytes had been transfected (estimated by RFP signal).

Single Myotube Ca²⁺ Imaging Experiments and Measurements of Myotube Diameter

The myotubes were loaded with 5 μm fluo-4 (fura-2 for measurements of resting myoplasmic Ca²⁺ levels) in an imaging solution (125 mm NaCl, 5 mm KCl, 2 mm KH₂PO₄, 2 mm CaCl₂, 25 mm HEPES, 6 mm glucose, 1.2 mm MgSO₄, 0.05% BSA (fraction V), pH 7.4) at 37 °C for 45 min. The myotubes were then washed three times with 200 μl of the imaging solution. The myotubes were transferred to an inverted stage microscope equipped with a Nikon ×40 oil-immersion objective (NA 1.30) (ECLIPSE Ti, Nikon). Before starting the single myotube Ca²⁺ imaging experiments, images of myotubes were captured for size comparison. Only GFP-positive myotubes were considered for size comparison. For the Ca²⁺ imaging experiments, images of the myotubes were captured using a high-speed monochromator with a 75-watt Xenon lamp (FSM150Xe, Bentham Instruments, Ltd.) as the light source and a 12-bit CCD camera (DVC-340M-OO-CL, Digital Video Camera Co.). Fluo-4 in the myotubes was excited at 494 nm, and fluorescence emission was measured at 516 nm. Data were displayed and analyzed using InCyt Im1 image acquisition and analysis software (version 5.29, Intracellular Imaging Inc.). Fura-2 in the myotubes was excited at 340 and 380 nm, and fluorescence emission was measured at 510 nm using High Speed InCyt Im2 image acquisition and analysis software (version 5.29, Intracellular Imaging Inc.). Ca²⁺ concentrations were calculated as described by Grynkiewicz et al. (31) using 225 nm as the Ca²⁺-fura-2 dissociation constant. KCl or caffeine was dissolved in the imaging solution and applied via an autoperfusion system with an 8-channel perfusion pipette (AutoMate Scientific). Cyclopiazonic acid (CPA) was dissolved in Me₂SO (<0.05%) and manually applied to the myotubes. Me₂SO (0.05%) alone had no effect on Ca²⁺ transients.

For SOCE experiments, myotubes loaded with fluo-4 were incubated with the imaging solution with zero Ca²⁺ for 5 min and then treated with CPA to induce SR Ca²⁺ depletion. Once the myoplasmic Ca²⁺ level returned to the baseline level, the imaging solution with 2 mm Ca²⁺ was added to the myotubes. To analyze Ca²⁺ transients obtained from Ca²⁺ imaging experiments, both the peak amplitude (maximum amplitude of a given peak, Table 1) and area (data not shown), which exhibited similar increases or decreases, were considered. For relatively long-term Ca²⁺ transients in CPA treatments or SOCE experiments (more than 20 min of recording), the peak areas were analyzed and the results are presented in Table 1.

TABLE 1.

Properties of Y141H-expressing myotubes

Values were normalized to the mean value of those from myotubes expressing wild-type JP2. Values are represented as the mean ± S.E. for the number of myotubes shown in parentheses. Values for the resting myoplasmic Ca²⁺ level, and KCl, caffeine, and CPA responses of wild-type and untransfected myotubes were adopted from our previous report (29).

* Significant difference versus wild-type (p < 0.05).

Mn²⁺ Quench Experiments

Fura-2-loaded myotubes were incubated with the imaging solution with zero Ca²⁺ for 2 min, and then Mn²⁺-containing solution (125 mm NaCl, 5 mm KCl, 2 mm KH₂PO₄, 25 mm HEPES, 6 mm glucose, 2 mm MnCl₂, 0.5 mm EGTA, 1.2 mm MgSO₄, 0.05% BSA (fraction V), pH 7.4) was applied to the myotubes for 5 min. The decrease in fura-2 fluorescence was monitored and the fluorescence quench rate was calculated by linear regression (y = a + bx) using Origin version 7. For linear regression, linear and stable traces (20 s after the exposure to Mn²⁺-containing solution) were used. The slopes derived from the linear regression were expressed as fluorescence arbitrary units per second.

Orai1 Knockdown by Small Interference RNA (siRNA)

Sequences of different siRNAs for Orai1 (GenBank^TM accession number NM_175423.3) were selected using a siRNA design software, siDirect (32), and are summarized in Table 2. The siRNAs were synthesized by Genolution (Seoul, Korea), and a scrambled siRNA was used as a negative control. Premature myotubes transfected with cDNA encoding Y141H were transfected with one of the siRNAs on differentiation day 4 in a mixture containing 600 μl of low-glucose DMEM, 60 μl of a transfection reagent (X-tremeGENE, Roche Diagnostics), and 200 nm of the specific synthetic siRNAs for 3 h according to the manufacturer's protocols (Roche and Genolution). After the transfection, fully differentiated myotubes on differentiation day 5 were subjected to further experiments. The transfection efficiency assessed by a cell viability assay using a siRNA for a homeogene supplied by Genolution was ∼95%. Optimal quantities of X-tremeGENE and siRNA and transfection frequencies were defined by evaluating the strongest effect of siRNAs.

TABLE 2.

Sequences of siRNAs used for knocking down Oria1 and primers used for qPCRs

Reverse Transcription and Quantitative Real-time PCR (RT-PCR and qPCR)

Total RNA was isolated from myotubes using QIAshredder and the RNeasy Mini Kit (Qiagen) according to the manufacturer's protocols. RT-PCR and qPCR were performed on the SmartCycler System (Cephid) using a One-Step kit (TaKaRa). Briefly, the following reaction mixture was prepared on ice: 12.5 μl of 2× One-Step SYBR RT-PCR buffer III, 2.5 units of Ex TaqHS, 0.5 μl of PrimeScript RT Enzyme Mix II, 0.2 μm of each PCR forward or reverse primer, and 50 ng of total RNA (25 μl of total reaction volume). Reaction conditions were 1 cycle of 42 °C for 5 min and 95 °C for 10 s for RT-PCR, 50 cycles of 95 °C for 5 s and 60 °C for 20 s for qPCR, and incubation at 60 °C for 1 min for the dissociation step. After the reaction was completed, signal verification was conducted with the amplification and melting curves using a comparative threshold cycle method from the SmartCycler System (Cephid). A mean quantity was calculated from duplicate or triplicate qPCR for each sample, and this quantity was normalized to the endogenous control gene, β-actin. Primers used for the qPCR are described in Table 2.

Immunocytochemistry

Myotubes were fixed in cold methanol (−20 °C) for 15 min and permeabilized with 0.05% Tween 20 in phosphate-buffered saline (PBS) for 1 min. After blocking myoblasts with 2% normal goat serum (Jackson ImmunoResearch) in PBS, the myoblasts were incubated with anti-STIM1 or anti-Orai1 antibody (10 μg/ml) for 3 h at room temperature, washed with 2% normal goat serum in PBS for 10 min, incubated with a Cy-3-conjugated anti-mouse (1:500, Jackson ImmunoResearch) or FITC-conjugated anti-rabbit secondary antibody (1:500, Sigma) for 45 min at room temperature, and visualized with an inverted fluorescence microscope using a ×40 objective lens (ECLIPSE Ti, Nikon) equipped with a monochrome camera (ProgRes MF, JENOPTIK Optical Systems, Inc.).

Preparation of Cell Lysate and Immunoprecipitation Assay

Myotubes or HL-1 cardiomyocytes were solubilized in a lysis buffer (1% Triton X-100, 10 mm Tris-HCl (pH 7.4), 1 mm Na₃VO₄, 10% glycerol, 150 mm NaCl, 5 mm EDTA, and protease inhibitor mixture (Roche Diagnostics)) overnight at 4 °C with gentle mixing (300 μl of the lysis buffer to the myotubes in a 10-cm dish). The solubilized lysate was then centrifuged at 1,500 × g for 30 s to remove insoluble matter. For the immunoprecipitation assay, the solubilized myotube lysate (800 μg of total protein) was incubated with anti-phosphoserine antibody (20 μg/mg, Chemicon International) overnight at 4 °C, followed by incubation with Protein G-Sepharose beads (Amersham Biosciences) at 4 °C for 4 h. The beads were washed five times with PBS. Proteins bound to the beads were eluted by boiling in a SDS sample buffer and subjected to immunoblot assay.

Immunoblot Assay

Myotube or HL-1 cardiomyocyte lysate (30 μg of proteins), or immunoprecipitants were subjected to 8 or 10% SDS-polyacrylamide gel electrophoresis, and the proteins on the gel were stained with Coomassie Brilliant Blue or transferred onto a polyvinylidene fluoride membrane at 100 V for 2 h. The membrane was blocked by 5% nonfat milk dissolved in PBS for 1 h, incubated with one of the primary antibodies overnight at 4 °C, washed three times with PBS containing 0.1% Tween 20 (PBST), and then incubated with one of the corresponding horseradish peroxidase-conjugated anti-mouse, anti-rabbit, or anti-donkey secondary antibodies (1:2,000, Jackson ImmunoResearch) for 45 min at room temperature. The membrane was washed three times with PBST and developed using SuperSignal ultrachemiluminescent substrate (Pierce). An anti-DHPR antibody (against α1 subunit of DHPR, 1:1,000) was obtained from Affinity BioReagents; an anti-RyR1 antibody (1:2,000) was obtained from Drs. J. Airey and J. Sutko (Developmental Studies Hybridoma Bank); anti-Orai1 and anti-STIM1 antibodies (1:500) were obtained from Abcam; and anti-JP2 (1:1,000), anti-TRPC1 (1:800), anti-TRPC3 (1:800), anti-TRPC4 (1:800), anti-TRPC6 (1:800), anti-PKC (1:500), anti-CaMK II (1:1,000), anti-CnA (1:1,000), and anti-α-actin (1:1,000) antibodies were obtained from Santa Cruz Biotechnology.

Cell Migration Assay

Cell migration assay was performed as previously described (33). Briefly, premature myotubes transfected with cDNA encoding wild-type JP2 or Y141H were scraped off with a pipette tip to make a long and thin acellular area. Images of the acellular area with the adjacent premature myotubes surrounding it were captured. After 12 h of scraping, images of the same acellular area were captured to evaluate the migration of premature myotubes to the acellular area.

Transmission Electron Microscopy (TEM)

For TEM observations, myotubes were fixed in a fixative solution (4% paraformaldehyde and 2.5% glutaraldehyde in 0.1 m phosphate buffer, pH 7.4) overnight at 4 °C. The fixed myotubes were washed with the phosphate buffer and post-fixed with 1% osmium tetroxide in phosphate buffer for 1 h at room temperature. The myotubes were dehydrated with ethyl alcohol and acetone, and embedded in epoxy resin (Epon 812). Ultra-thin sections (70–80 nm) were obtained using an ultramicrotome (Ultracut UCT ultramicrotome, Leica). The sections were double-stained with uranyl acetate and lead citrate, and examined under a transmission electron microscope (JEM1010, JEOL Ltd.) at 60 kV.

Statistical Analysis

Results are presented as the mean ± S.E. for the number of myotubes shown in parentheses in Table 1. Values were normalized to the mean value of those from corresponding controls. Significant differences were analyzed using a paired t test (GraphPad InStat, version 2.04). Differences were considered to be significant at p < 0.05. Graphs were prepared using Origin version 7.

RESULTS

Y141H Mutation of JP2 Induces Hypertrophy in Skeletal Myotubes

Tyr¹⁴¹ in JP2 is one of the residues comprising the first group of MORN motifs (Fig. 1A). A human HCM-associated JP2 mutant, Y141H, was expressed in mouse primary skeletal myotubes as described under “Experimental Procedures.” Myotube lysate from the plates transfected with cDNA encoding either wild-type JP2 or Y141H showed approximately twice as much JP2 protein as myotube lysate from the untransfected plates (Fig. 1B). Considering the heterogeneous nature of transient transfection and the ∼40% transfection efficiency (estimated by the RFP signals, Fig. 1C), an average amount of total JP2 proteins (both endogenous and exogenous) in the transfected myotubes would be 3.5-fold compared with that in untransfected myotubes. Interestingly, a significant increase in diameter was found in Y141H-expressing myotubes compared with wild-type controls (increased by 1.53 ± 0.13-fold) (Fig. 1D and Table 1). These results indicate that a single mutation in JP2 at Tyr¹⁴¹ induced hypertrophy in skeletal myotubes, similar to the hypertrophy observed in the cardiac muscle from HCM patients.

FIGURE 1.

An increase in the diameter of Y141H-expressing myotubes. A, schematic diagram of mouse JP2 primary sequence. There are 8 MORN motifs (shown in dark gray) and a TM domain (shown in light gray). The location of Tyr¹⁴¹ is indicated. B, myotube lysate from the plates transfected with either wild-type JP2 or Y141H was subjected to immunoblot assay with anti-JP2 antibody and showed approximately twice as much JP2 protein as myotube lysate from the untransfected plates (considering ∼40% transfection efficiency, an average 3.5-fold increase in the transfected myotubes). α-Actin was used as a loading control. C, transfected myotubes with either wild-type JP2 or Y141H were visualized by the presence of the RFP signal. Diameters of the thickest part of the given myotubes were measured and representative diameters of myotubes are indicated by white arrows. Bar represents 200 μm. D, histograms are shown for the distribution of diameters normalized to the mean value of diameters from wild-type controls. Lines indicate the maximum or minimum values. Boxes indicate mean ± S.E. Squares in boxes indicate the mean value. Y141H-expressing myotubes showed an increase in diameter. *, significant difference compared with wild-type (p < 0.05).

Reduced Gain of EC Coupling and Abnormal JMC in Y141H-expressing Skeletal Myotubes

To measure myoplasmic Ca²⁺ transients per unit area during EC coupling, KCl (inducing membrane depolarization) was applied to Y141H-expressing myotubes. Y141H-expressing myotubes showed a significant decrease in myoplasmic Ca²⁺ transients with no change in the activation slope compared with wild-type controls (Fig. 2A and Table 1). Because Ca²⁺ transients would be expected to be larger if the size of the myotubes increased, the net decrease of Ca²⁺ transients during EC coupling in Y141H-expressing myotubes might possibly be larger than that presented in Fig. 2A. The expression level of the α1 subunit of DHPR (the membrane voltage-sensing and Ca²⁺-conducting pore-forming subunit of DHPR) or RyR1 was not significantly changed (Fig. 5B), suggesting that the decrease in myoplasmic Ca²⁺ transients during EC coupling was not simply due to a reduction in DHPR or RyR1 expression.

FIGURE 2.

Decreased gain of EC coupling in Y141H-expressing myotubes. KCl (A) or caffeine (B) was applied to myotubes loaded with fluo-4. Activation slopes were fitted using linear equation and are represented by dotted lines. For comparison, vertical lines are shown as solid lines. Histograms are shown for normalized peak amplitude (left) or activation slope (right) to the mean value of those from wild-type controls. Y141H-expressing myotubes showed a significant decrease in myoplasmic Ca²⁺ transients per unit area in response to KCl but not to caffeine. C, SR Ca²⁺ in myotubes loaded with fluo-4 was depleted by treatment with 10 μm CPA. Ca²⁺ transients per unit area induced by CPA treatments were summarized as histograms (the peak area was normalized to the mean value of those from wild-type controls). Small peaks at the moment of CPA treatment were artifacts of manual CPA treatments. There was no change in the SR Ca²⁺ level of Y141H-expressing myotubes. *, significant difference compared with wild-type (p < 0.05).

FIGURE 5.

An increase in Orai1 expression in Y141H-expressing myotubes and HL-1 atrial cardiomyocytes. Each myotube lysate was subjected to immunoblot assay with one of the antibodies against Ca²⁺-dependent protein kinase or phosphatase (A), or one of various Ca²⁺ channels (B). α-Actin was used as a loading control. PKC, protein kinase C; CaMK II, calmodulin-dependent protein kinase II; CnA, calcineurin A. Y141H-expressing myotubes showed increases in PKC and Orai1 expression compared with wild-type controls (marked by asterisks, considering ∼40% transfection efficiency, average 2- and 4.8-fold increases, respectively). HL-1 cardiomyocytes were transfected with cDNA encoding wild-type JP2 or Y141H. Y141H-expressing HL-1 cardiomyocytes also showed increases in both Orai1 expression (marked by an asterisk, considering ∼50% transfection efficiency, an average 1.8-fold increase) (C) and cell size (D) compared with wild-type controls. α-Actin was used as a loading control. Boxed areas in Y141H were enlarged in the left-hand panels in D. Bar represents 50 μm.

There are two possible explanations for the reduced gain of EC coupling in the Y141H-expressing myotubes. 1) RyR1 activity might be reduced and result in a reduced gain in EC coupling, or 2) the spatial arrangement of RyR1 and DHPR might not be complete enough to induce the full gain of EC coupling. To test the first possibility, we measured myoplasmic Ca²⁺ transients per unit area in response to a direct RyR1 agonist, caffeine. Y141H-expressing myotubes showed no significant change in myoplasmic Ca²⁺ transient in response to caffeine, and the activation slope of the Ca²⁺ transients was comparable with that of wild-type controls (Fig. 2B and Table 1), suggesting that the reduced gain of EC coupling was not due to a change in RyR1 activity. In accordance with the unchanged RyR1 activity, there was also no significant change in the amount of SR-stored Ca²⁺ (estimated by treatment with 10 μm CPA, an inhibitor of the SR Ca²⁺ uptake pump) (Fig. 2C).

To test the latter possibility, myotubes-expressing wild-type JP2 or Y141H were examined by TEM. Unlike well organized JMC in wild-type controls, it is of interest that abnormal JMC (multilayered tubule structures) was frequently found between two SRs or near the SR in Y141H-expressing myotubes (Fig. 3, right, indicated by arrowheads). Considering that paired or close arrangements of RyRs and DHPR on JMC are a necessary factor for authentic skeletal or cardiac EC coupling, respectively, the multilayered abnormal tubule structures could be one of the causes for the reduced gain of EC coupling in Y141H-expressing myotubes. Hypertrophy, a reduced gain of EC coupling, and abnormal JMC are dominant-negative effects of the Y141H mutant over the native JP2, because native JP2 is endogenously expressed in skeletal muscle.

FIGURE 3.

Abnormal JMC in Y141H-expressing myotubes. Myotubes-expressing wild-type or Y141H were visualized using TEM observations. Areas in the numbered boxes were enlarged. In Y141H, the images in boxes 2 and 3 are considered as 90° horizontally rotated and tilted images of box 1, respectively. Multilayered tubule structures near the SR membranes (indicated by white arrowheads) were frequently observed in Y141H-expressing myotubes. T in wild-type indicates authentic t-tubule structures. Bars represent 2 μm.

Increase in Resting Myoplasmic Ca²⁺ Levels in Y141H-expressing Skeletal Myotubes

To determine the molecular mechanism(s) underlying the reduced gain of EC coupling in Y141H-expressing myotubes, resting myoplasmic Ca²⁺ levels were examined. Y141H-expressing myotubes maintained a significantly higher resting myoplasmic Ca²⁺ level per unit area than wild-type controls (Fig. 4A and Table 1, an increase of ∼70%). The absolute amount of total Ca²⁺ ions in the myoplasm of a single Y141H-expressing myotube is supposed to be much greater than that of wild-type controls. Because Y141H-expressing myotubes were hypertrophied, the absolute myoplasmic volume of the hypertrophic myotubes (in terms of three dimensions) was much greater than that of wild-type controls.

FIGURE 4.

An increase in resting myoplasmic Ca²⁺ level in Y141H-expressing myotubes. A, resting myoplasmic Ca²⁺ levels per unit area in myotubes loaded with fura-2 were measured. *, significant difference compared with wild-type (p < 0.05). The resting myoplasmic Ca²⁺ level was increased in Y141H-expressing myotubes. B, Ca²⁺ entry at rest per unit area was measured using the Mn²⁺ quench experiments. The arrow shows the time point when the Mn²⁺-containing solution was applied to myotubes. The rate of fluorescence decay for each trace was obtained from the slope of linear regression and presented by histograms (right). Y141H-expressing myotubes showed no significant change in Mn²⁺ quench rate compared with wild-type controls.

To find possible upstream or downstream factors for the elevation of the resting myoplasmic Ca²⁺ level in Y141H-expressing myotubes, first, Ca²⁺ entry at rest was measured using Mn²⁺ quench experiments. The decay rate in fura-2 fluorescence in Y141H-expressing myotubes after Mn²⁺ exposure was decreased very little but not significantly different from that of the wild-type controls (Fig. 4B), suggesting that a change in Ca²⁺ entry at rest was not a reason for the increase in the resting myoplasmic Ca²⁺ level. Second, the expression of Ca²⁺-dependent protein kinases or phosphatases, which are known to be involved in HCMs, was examined by immunoblot assay. PKC expression was significantly increased in Y141H-expressing myotubes compared with wild-type controls (Fig. 5A). Therefore, it is likely that the reduced gain of EC coupling in Y141H-expressing myotubes would result from change(s) in PKC-mediated signaling.

On the other hand, Tyr¹⁴¹ itself is an irrelevant mutant to phosphorylation by PKC, but it is possible that the mutation at Tyr¹⁴¹, which is an adjacent residue to Ser¹⁶⁵ (the substantial and only phosphorylation site of JP2 by PKC, supplemental Fig. S1), could mask the phosphorylation at Ser¹⁶⁵ by blocking the accessibility of PKC to Ser¹⁶⁵. Immunoprecipitation of myotube lysate with anti-phosphoserine antibody, followed by immunoblot assay of the immunoprecipitates with anti-JP2 antibody showed that the total amount of phosphorylated JP2 proteins at the serine residue was not altered by the expression of Y141H (supplemental Fig. S2). Therefore, it seems that Y141H was not directly affected by the increased PKC expression.

An Increase in SOCE Mediated by Orai1 in Y141H-expressing Skeletal Myotubes

Third, the expression level of Ca²⁺ conducting channels was examined by immunoblot assay using myotube lysate from Y141H-expressing myotubes (Fig. 5B). As mentioned earlier, the expression levels of the two major EC coupling-mediating Ca²⁺ channels (DHPR and RyR1) were not changed. The expression of five extracellular Ca²⁺ entry channels that are known to be expressed in skeletal muscle (TRPC1, TRPC3, TRPC4, TRPC6, and Orai1) was also examined. Among them, interestingly, Orai1 (a major protein responsible for SOCE in response to SR Ca²⁺-depletion in skeletal muscle) was significantly up-regulated (∼4.8-fold considering 40% transfection efficiency) with no change in the expression level of its functional partner, STIM1 (a Ca²⁺-sensing protein on SR membrane). As shown in Y141H-expressing myotubes, Orai1 expression was also increased in Y141H-expressing HL-1 atrial cardiomyocytes (Fig. 5C) as well as cell size (i.e. hypertrophy, Fig. 5D).

To test whether the increased Orai1 expression also affected an Orai1-mediated functional event (i.e. SOCE), the SR of Y141H-expressing myotubes was depleted by treatment with CPA in the absence of extracellular Ca²⁺, and then 2 mm Ca²⁺ was applied to the extracellular bath to induce SOCE. As expected, SOCE was significantly increased (Fig. 6A and Table 1, an increase of ∼88%). In addition, puncta formed by oligomerized STIM1 and Orai1 (the necessary organization for SOCE) were also more frequently found in Y141H-expressing myotubes (Fig. 6B). Therefore, increased SOCE due to the increased expression of Orai1 could be one reason for the elevated resting myoplasmic Ca²⁺ level in Y141H-expressing myotubes.

FIGURE 6.

Increases in both SOCE and puncta in Y141H-expressing myotubes. A, myotubes loaded with fluo-4 were depleted by treatment of CPA in the absence of Ca²⁺ in the imaging solution, and 2 mm Ca²⁺ was added to the extracellular bath to induce SOCE. A representative trace for each group is presented. Y141H-expressing myotubes showed a significantly increased SOCE. SOCE per unit area was summarized as histograms (the peak area was normalized to the mean value of those from wild-type controls). *, significant difference compared with wild-type (p < 0.05). B, myotubes were double-stained with anti-Orai1 and anti-STIM1 antibodies. Merged images show more puncta in Y141H-expressing myotubes. Boxed areas in the merged images were enlarged in the right-hand panels. DIC, differential interference contrast microscopy.

To confirm that the increase in size in Y141H-expressing myotubes was mediated by the increased Orai1 expression, Orai1 was knocked down in Y141H-expressing myotubes. To interfere with the mRNA of Orai1, nine different siRNAs for Orai1 were designed (shown in Table 2) and subjected to transfections to Y141H-expressing myotubes as described under “Experimental Procedures.” Real-time qPCR analysis of cDNA from each transfected myotube batch showed that number 4 siRNA was the most efficient in knocking down Orai1 (77.5 ± 1.5% reduction in mRNA and ∼94% reduction in protein, Fig. 7, A and B, and supplemental Fig. S3). Therefore, number 4 siRNA was subjected to further experiments. Premature myotubes transfected with cDNA encoding Y141H were transfected with number 4 siRNA on differentiation day 4. Transfection efficiency for Y141H was visualized by the presence of RFP signals (Fig. 7C, right-hand panel), and transfection efficiency for siRNA was assessed by a cell viability assay using a siRNA for a homeogene described under “Experimental Procedures.” Y141H-expressing and Orai1 knockdown myotubes showed no significant change in diameter (Fig. 7C, white arrows in the right-hand panel) and resting myoplasmic Ca²⁺ level (Fig. 7D) compared with either untransfected control or scrambled siRNA- or number 4 siRNA-transfected control. Therefore, the increased Orai1 expression resulted in increases in both size and resting myoplasmic Ca²⁺ levels in Y141H-expressing myotubes.

FIGURE 7.

No hypertrophic phenomenon in Y141H-expressing myotubes by knocking down Orai1. To interfere with the mRNA of Orai1, myotubes were transfected with the siRNAs for Orai1 or scrambled control as described under “Experimental Procedures.” A, qPCR analysis showed that Orai1 mRNA was reduced by 77.5 ± 1.5% by number 4 siRNA transfection compared with untransfected or scrambled controls. The data are the mean ± S.E. of duplicate independent experiments. Significant reduction in the mRNA level of Orai1 is indicated by a white asterisk. For the results of other siRNAs, see Table 2 and supplemental Fig. S3. B, myotube lysate from Y141H-expressing and Orai1 knockdown myotubes was subjected to immunoblot analysis with anti-Orai1 antibody. The expression of the Orai1 protein was reduced by ∼94%. C, number 4 siRNA was transfected to Y141H-expressing myotubes. Y141H expression in the Y141H-expressing and Orai1 knockdown myotubes was visualized by the presence of RFP signal (bottom panel). Diameters of the thickest part of the given myotubes were measured and representative diameters of myotubes are indicated by white arrows (left-hand panel). Y141H-expressing and Orai1 knockdown myotubes showed no significant change in diameter compared with either untransfected control or scrambled siRNA- or number 4 siRNA-transfected control. Bar represents 300 μm. DIC, differential interference contrast microscopy. D, resting myoplasmic Ca²⁺ levels per unit area in Y141H-expressing and Orai1 knockdown myotubes loaded with fura-2 were measured. Resting myoplasmic Ca²⁺ level was not significantly changed compared with either untransfected control or scrambled siRNA- or number 4 siRNA-transfected control (colored in gray).

DISCUSSION

Hypertrophy in Skeletal Myotubes Caused by Two Human HCM-associated JP2 mutants, Y141H and S165F, Involves Altered Ca²⁺ Signaling via Different Proteins

In summary, we examined the effect of the Y141H mutant of JP2 on skeletal muscle. Consistent with the S165F mutant of JP2 (29), a dominant-negative inhibition by Y141H over the native JP2 resulted in several consequences, including significant myotube hypertrophy, a decreased gain of EC coupling, and an increase in the resting myoplasmic Ca²⁺ level. Different from S165F, Y141H-expressing myotubes involved two new characteristics: abnormal JMC and an increase in SOCE via Orai1.

It is well known that various hypertrophic stimuli elevate intracellular Ca²⁺ levels, which mediate the progression of cardiac hypertrophy via various routes: for example, activation of Ca²⁺-related proteins (34). In accordance with this, the expression of Y141H in skeletal myotubes induced hypertrophy with a reduced gain of EC coupling and an elevated resting myoplasmic Ca²⁺ level. However, to reach the hypertrophic phenomena, a different subroute utilizing different proteins was involved in Y141H-expressing myotubes compared with S165F. First, unlike S165F, which is an unphosphorylable mutant by PKC (29), the phosphorylation level of Y141H by PKC was about the same as that of wild-type JP2 (supplemental Fig. S2). Second, Y141H-expressing myotubes showed an increase in Orai1 expression accompanying increases in both physiological SOCE and morphological puncta formation. However, there was no change in Orai1 expression in S165F-expressing myotubes (supplemental Fig. S4), suggesting that Orai1 is a skeletal hypertrophy-mediating protein by Y141H but not by S165F. Interestingly, Orai1 seems to also be a hypertrophy-mediating protein in cardiac muscle, because Orai1 knock-out in neonatal rat ventricular cardiomyocytes diminished cell size and even completely blocked phenylephrine-mediated hypertrophy (35). In addition, in the present study, HL-1 atrial cardiomyocytes-expressing Y141H showed increases in both Orai1 expression and cell size.

On the other hand, PKC has reportedly suppressed SOCE by the phosphorylation of Orai1 in HEK293 cells (36). In the present study, the increased expression of PKC in Y141H-expressing myotubes may be a compensatory action for the increased Orai1 expression.

Tyr¹⁴¹ and Ser¹⁶⁵ Residues of JP2 Play Different Roles in Ca²⁺ Signaling of Skeletal Muscle

Tyr¹⁴¹ in JP2 is one of the residues comprising MORN motifs that interact with t-tubule membranes to “structurally” form JMC, and JMC allows EC coupling-mediating and -regulating proteins to be positioned close enough for functionally efficient cardiac and skeletal EC coupling (2, 12, 13). In the present study, a mutation at Tyr¹⁴¹ substantially induced abnormal JMC and possibly subsequent reduction in the gain of EC coupling in skeletal myotubes. On the other hand, Ser¹⁶⁵ of JP2 is an irrelevant residue to MORN motifs by locating in the joining region between two groups of MORN motifs (supplemental Fig. S1). Secondary structure prediction for JP2 using a program, Jpred3 Incorporating Jnet (37), suggests that the region containing Ser¹⁶⁵ forms a short α-helix (151 to 166 residues) (21). The region covering the short α-helix confers TRPC3-binding abilities on JP2 (20, 21). A single mutation at Ser¹⁶⁵ greatly diminishes the binding ability of JP2 to TRPC3 (29). Taken together, it seems that Tyr¹⁴¹ participates in the formation of complete JMC as a building block (structural integrity for Ca²⁺ signaling), and Ser¹⁶⁵ is one of the tuning buttons for complete connections among EC coupling-mediating and -regulating proteins on JMC (functional integrity for Ca²⁺ signaling). Therefore, it seems that Tyr¹⁴¹ and Ser¹⁶⁵ of JP2 play different roles in the Ca²⁺ signaling of skeletal muscle, and the location of mutations in JP2 could be a code for discriminating among several subroutes of hypertrophic pathways in skeletal muscle.

Authentic JMC in the Presence of JP2 Are Key Structures for Striated Muscle Function

Y141H-expressing myotubes showed abnormally multilayered tubule structures near the SR instead of authentic JMC. Under this circumstance, the rate of physical coupling between DHPR and RyR1 could be decreased (more orphaned RyRs unoccupied by DHPR) due to the widespread distribution of the limited numbers of DHPRs on the wider tubule membranes. The possibility for the presence of more orphaned RyRs in Y141H-expressing myotubes was strongly supported by the reduced gain of EC coupling unaccompanied by a change in RyR1 activity, in DHPR and RyR1 expression, and in SR Ca²⁺ content.

In the case of cardiac muscle cells, the efficiency of EC coupling would also be lowered due to abnormally multilayered tubule structures that pose physical obstacles to the diffusion of Ca²⁺ ions to RyR2. Indeed, a reduced gain of EC coupling due to dispersion of JMC without altered Ca²⁺ channel expression or SR Ca²⁺ content (like Y141H-expressing myotubes in the present study) was reported in cardiomyocytes from heart failure or cardiac hypertrophic rat models and from a ventricular arrhythmia mouse model (38–40). From the viewpoint of JP2, a gradual reduction in JP2 expression and contractility was found in ventricular myocytes from the early hypertrophic rat (26), in hypertrophic and dilated cardiomyopathic mouse models (24), and in a pressure overload-induced cardiac hypertrophic rat model (25). Clinically, cardiac tissue from HCM patients showed a reduction in JP2 expression, and JP2 knockdown in HL-1 cardiomyocytes resulted in hypertrophy with reduced gain of EC coupling (41). Therefore, in both skeletal and cardiac muscles, abnormal JMC due to the reduced expression of native JP2 or the presence of mutated JP2s could be an ultrastructural substrate for abnormal muscle functions.

The Hypertrophy in Y141H-expressing Myotubes Could Be Related to Faster Migration of Premature Myotubes, Possibly Due to Increased SOCE

Migration and fusion (the first phase of fusion) of myoblasts are key steps in the myogenesis and regeneration of skeletal muscle. To form long and multinucleated myotubes during skeletal muscle differentiation, myoblasts migrate in all directions and form pre-fusion alignment before actual fusion (42). The second phase of fusion, which is called maturation in normal physiology or hypertrophy in above normal physiology, permits premature myoblasts to become longer and thicker. It is interesting that more fusion was observed in Y141H-expressing myotubes than wild-type controls during differentiation under an electron microscope, which was not previously observed in S165F-expressing myotubes. To find the possible reason(s) for this result, migrations of premature myotubes were examined on differentiation day 3, when premature myotubes vigorously migrate to form longer and thicker mature myotubes. Y141H-expressing premature myotubes migrated to an acellular area faster than wild-type controls (supplemental Fig. S5). Vascular smooth muscle cells with a reduced SOCE by the knockdown of STIM1 and Orai1 showed the inhibition of cell migration (43). C2C12 cells (a mouse myoblast cell line) with a reduced SOCE by TRPC1 knockdown showed a slow down in myotube formation (33). In the present study, Y141H-expressing myotubes showed increases in both Orai1 expression and SOCE. According to previous reports and our results in the present study, the increased SOCE seems to be related to higher migration efficiency and hypertrophy in Y141H-expressing myotubes.

Orai1 and SOCE Are Effectors of Mutated JP2-bearing Striated Muscle Hypertrophy

As addressed earlier, STIM1 is a sensor protein for endoplasmic reticulum/SR Ca²⁺ depletion (5, 6, 44). Ca²⁺ depletion causes STIM1 to be oligomerized and redistributed to the endoplasmic reticulum/SR membrane near the cellular periphery, and to interact with Orai1, which is called puncta (a prerequisite for SOCE via Orai1) (5, 6, 9, 10, 44). In particular, puncta in skeletal muscle were pre-assembled naturally during the differentiation of myoblasts to myotubes (10). In the present study, fully differentiated mouse primary skeletal myotubes also showed pre-assembled puncta (ready-to-go forms, which is a prerequisite for SOCE via Orai1 but not the necessary and sufficient condition for SOCE). Skeletal and cardiac muscles seem to take advantage of their specialized JMC (triad or diad) to achieve the pre-assembled puncta, and these ready-to-go puncta are compatible with the rapid activation of SOCE in rat skeletal muscle fibers (activation in less than 1 s (45)). Y141H-expressing myotubes showed more preassembled puncta than wild-type controls, and there are two potential mechanisms for this. One is that the abnormally multilayered tubule structure in Y141H-expressing myotubes may increase the chance of a juxtaposed arrangement of tubules and SR membranes, which favors preassembly of the puncta. The other is that the increased Orai1 expression in Y141H-expressing myotubes might act as a puncta-tropic factor through mass action: a quantitative gain increases qualitative gain. In either case, more preassembled puncta and subsequently more SOCE could contribute to the elevation of the resting myoplasmic Ca²⁺ level in Y141H-expressing myotubes. If this is the case, Orai1 and SOCE could be effectors of hypertrophic phenomena in striated muscle.

In vascular smooth muscle and HEK293 cells, STIM1 strongly suppresses Ca_V1.2, while activating Orai1 (a reciprocal control of Ca_V1.2 and Orai1 by STIM1 at the same time) (46). A preponderance of Orai1 might induce more STIM1-Orai1 interaction (i.e. more SOCE) and more suppression of DHPR (Ca_V1.1, the skeletal isoform of Ca_V1.2, i.e. less EC coupling) as shown in Y141H-expressing myotubes. It is, therefore, possible that another type of retrograde signaling from the SR to the t-tubule membrane via STIM1 would exist in striated muscle.

Overall, JP2 has recently emerged as a hypertrophy-related protein by studies on HCM patients and various hypertrophic animal models. In the present study, we suggest how the Y141H mutant of JP2 is involved in skeletal muscle hypertrophy at the cellular level, and that an increase in SOCE via Orai1 could be a novel factor in the increased myoplasmic Ca²⁺ levels of JP2-related muscle hypertrophy. It is not currently clear whether the JP2 mutation plays a primary or secondary role in the development of hypertrophy. However, the present study suggests that JP2 could be a new molecular target for the regulation of abnormal Ca²⁺ homeostasis in striated muscle hypertrophy, and provides insight into possible varieties of subcellular mechanisms by which this mutant causes cardiac hypertrophy in humans bearing this mutation.